Abstract

Faithfull genome partitioning during cell division relies on the Spindle Assembly Checkpoint (SAC), a conserved signaling pathway that delays anaphase onset until all chromosomes are attached to spindle microtubules. Mps1 kinase is an upstream SAC regulator that promotes the assembly of an anaphase inhibitor through a sequential multi-target phosphorylation cascade. Thus, the SAC is highly responsive to Mps1, whose activity peaks in early mitosis as a result of its T-loop autophosphorylation. However, the mechanism controlling Mps1 inactivation once kinetochores attach to microtubules and the SAC is satisfied remains unknown. Here we show in vitro and in Drosophila that Protein Phosphatase 1 (PP1) inactivates Mps1 by dephosphorylating its T-loop. PP1-mediated dephosphorylation of Mps1 occurs at kinetochores and in the cytosol, and inactivation of both pools of Mps1 during metaphase is essential to ensure prompt and efficient SAC silencing. Overall, our findings uncover a mechanism of SAC inactivation required for timely mitotic exit.

The events that underlie SAC inactivation are less well understood. The phosphatases PP1-γ and PP2A-B56 appear to play an important role in this process by dephosphorylating KNL1-MELT motifs (London et al., 2012; Espert et al., 2014; Nijenhuis et al., 2014), which contributes to the removal of SAC proteins from kinetochores following microtubule attachments (Etemad and Kops, 2016). Furthermore, elegant in vitro studies demonstrated that binding of Mps1 to the calponin homology (CH) domains of Ndc80 and Nuf2 is inhibited by microtubules (Hiruma et al., 2015; Ji et al., 2015). Although this competition mechanism precludes the recruitment of Mps1 to bioriented kinetochores, it is however insufficient to remove all Mps1 from kinetochores even after the formation of stable end-on attachments (Aravamudhan et al., 2015; Hiruma et al., 2015; Ji et al., 2015). Work in budding yeast proposes that end-on attachments physically separate residual Mps1 from the Spc105/KNL1 phosphodomain, hence disrupting MELT phosphorylation at metaphase kinetochores (Aravamudhan et al., 2015). While this may contribute to silence the SAC in yeast, this model does not account for the highly dynamic behaviour of active Mps1 that has been observed in human cells (Jelluma et al., 2010). The association of Mps1 with the Ndc80 complex is very transient and the kinase diffuses rapidly into the cytoplasm, which is likely to allow Mps1 to overcome the increased spatial separation between its kinetochore receptor and Spc105/KNL1 MELT motifs. Moreover, kinetochore recruitment of Mad1-Mad2 in metazoa is also mediated through an Spc105/KNL1-independent pathway (Schittenhelm et al., 2009; Caldas et al., 2015; Silió et al., 2015). This is known to rely on the recruitment of the Rod/ZW10/Zwilch (RZZ) complex to unattached kinetochores, an event that is also controlled by Mps1 kinase (Santaguida et al., 2010). Importantly, in addition to its role at unattached kinetochores, Mps1 also contributes to SAC signaling in the cytoplasm. Soluble Mps1 is required for the assembly of pre-mitotic MCC that determines the peak time of anaphase onset in the absence of kinetochore–microtubule attachment problems (Rodriguez-Bravo et al., 2014). Moreover, Mps1 lacking its kinetochore-binding domain is sufficient to delay mitotic exit in mouse embryonic fibroblasts and RPE1 cells challenged with spindle poisons (Foijer et al., 2014; Maciejowski et al., 2010) and cytosolic Mps1 was shown to be required to support SAC arrest caused by kinetochore-tethered Mad1 (Maldonado and Kapoor, 2011). Therefore, in addition to the control exerted on Mps1 kinetochore recruitment and to alterations on kinetochore architecture, other mechanisms must contribute to disrupt SAC signaling upon stable end-on microtubule attachments. Here we show that in Drosophila, PP1-87B/PP1-γ dephosphorylates the T-loop of cytosolic and kinetochore-associated Mps1. This renders both pools of Mps1 inactive during metaphase, which we find to be critical for rapid SAC silencing and timely mitotic exit.

Results

Depletion of PP1-87B results in a metaphase delay with stable kinetochore-microtubule attachments

(A,B) Kymograph analysis of mitotic progression monitored by live-cell imaging (A) and corresponding quantification of Mad1-EGFP kinetochore levels (B) of control, PP1-87B depleted or Mps1 and PP1-87B co-depleted Drosophila S2 cells expressing mCherry-Tubulin and Mad1-EGFP under control of Mad1 promoter. The mean time from nuclear envelope breakdown to anaphase onset for each condition is displayed in the corresponding kymograph. Mad1-EGFP fluorescence intensities at kinetochores were corrected for cytosolic signal (N ≥ 9 cells for each condition). Vertical dashed line indicates the frame corresponding to nuclear envelope breakdown. Horizontal scale bar: 5 min. Vertical scale bar: 5 μm. (C,D) Representative immunofluorescence images (C) and corresponding quantifications (D) of inter-kinetochore distances and relative levels of Aurora B T232 phosphorylation (AurBT232Ph) at unaligned and aligned chromosomes in control and PP1-87B depleted S2 cells. AurBT232Ph relative levels were plotted over the inter-kinetochore distance measured as the distance between centroids of CID pairs. AurBT232Ph fluorescence intensities were determined relative to CID signal (N ≥ 58 kinetochore pairs for each condition). Scale bar: 5 μm. (E) Quantification of inter-kinetochore distances measured in chromosomes aligned at the metaphase plate of control and PP1-87B depleted S2 cells. Inter-kinetochore distances were measured as the distance between centroids of identified CID pairs (N ≥ 86 kinetochores for each condition). (F,G) Representative immunofluorescence images (F) and corresponding quantification (G) of cold-stable kinetochore fibers in control, PP1-87B and Ndc80 depleted S2 cells. CID immunolocalization was used as kinetochore reference. The insets display magnifications of the outlined regions. The graph represents the % of kinetochores attached to cold-stable microtubules per cell (N ≥ 10 cells for each condition). (H,I) Analysis of microtubules turnover rates by speckle contrast fadeout of GFP-α-Tubulin. (H) Contrast fadeout–time curves of GFP-α-Tubulin fluorescent speckles (lines) and their time point means (dots) measured in rectangular areas enclosing the spindle of control and PP1-87B-depleted S2 cells in prometaphase and metaphase. The rate of speckle contrast fadeout was calculated to obtain microtubule turnover rates. (I) Table showing microtubules half-lives of non-kinetochore- (non-KT-MTs) and kinetochore-microtubules (KT-MTs) of control and PP1-87B-depleted S2 cells determined by inducible speckle imaging in prometaphase and metaphase. The average speckle intensity squared-contrast at each time point was fit to a double-exponential curve A1*exp(-k1*t)+A2*exp(-k2*t), in which t is time, A1 represent the less stable population (non-KT-MTs) and A2 the more stable population (KT-MTs) with decay rates of k1 and k2, respectively. The turnover half-life for each population was calculated as ln2/k (N ≥ 7 metaphase cells for each condition). Data information: in (A), (B), (E), (G) and (H) data are presented as mean ± SD. Numerical source data for this figure are provided in Figure 1—source data 1.

The Video illustrates ISI imprinting and progressive contrast fade-out of GFP–α-tubulin in a control metaphase spindle. The observed speckle contrast fadeout of tubulin is due to microtubule turnover. The time between frames is variable and is presented on the corresponding excel source data file.

The Video illustrates ISI imprinting and progressive contrast fade-out of GFP–α-tubulin in a PP1-87B depleted metaphase spindle. The observed speckle contrast fadeout of tubulin is due to microtubule turnover. The time between frames is variable and is presented on the corresponding excel source data file.

PP1-87B antagonizes Mps1 T-loop autophosphorylation at kinetochores

The results described in the previous section suggest that the metaphase delay evoked by PP1-87B depletion results from problems in SAC silencing. Although, Mps1 is dispensable for kinetochore localization of Bub1 and BubR1 in flies (Schittenhelm et al., 2009; Conde et al., 2013) its activity is nonetheless required for Mad1 and Mad2 recruitment and therefore for SAC signaling (Althoff et al., 2012; Conde et al., 2013). In accordance, co-depletion of Mps1 with PP1-87B abolished Mad1 kinetochore recruitment and led to a premature anaphase onset, demonstrating that Mps1 is required for the SAC-dependent metaphase delay observed in PP1-87B RNAi (Figure 1A,B). Therefore, we sought to examine Mps1 kinetochore recruitment and activation in S2 cells depleted of PP1-87B. Immunofluorescence analysis showed that depletion of PP1-87B has no significant impact on Mps1 kinetochore localization. Mps1 accumulates at prometaphase kinetochores and its levels decrease significantly as microtubules attach, with a fraction of Mps1 persisting at kinetochores of metaphase chromosomes (Figure 2A–D). This residual pool of Mps1 is retained at bioriented kinetochores until anaphase onset, even after establishment of inter-kinetochore tension, as revealed by live imaging of S2 cells expressing EGFP-Mps1 under control of Mps1 cis-regulatory region (Figure 2—figure supplement 1A,B and Video 6). To evaluate Mps1 activation status we used a phospho-specific antibody that recognizes the conserved activating autophosphorylation (T490Ph) on Mps1 T-loop (Jelluma et al., 2008). Interestingly, we found that depletion of PP1-87B resulted in a substantial increase of Mps1 T-loop autophosphorylation during prometaphase and on metaphases kinetochores stably attached to microtubule bundles (Figure 2A–D, Figure 2—figure supplement 1C,D and Figure 2—figure supplement 2A–F with phospho-antibody characterization described in supplementary information). Moreover, depletion of PP1 noncatalytic subunit Sds22/PPP1R7, which positively regulates the phosphatase activity at kinetochores (Posch et al., 2010; Wurzenberger et al., 2012; Heroes et al., 2013), mimicked PP1-87B RNAi by increasing Mps1 T490 phosphorylation both at unattached and aligned kinetochores (Figure 2—figure supplement 3A–D). Mps1 association with prometaphase kinetochores in human cells is highly transient (Jelluma et al., 2010). Likewise, Drosophila Mps1 exhibits a fast turnover rate at unattached kinetochores that relies in part on its kinase activity, as revealed by FRAP measurements of wild type (WT) and kinase-dead (KD) versions of EGFP-Mps1 expressed in S2 cells (Figure 2E,F). In line with the observed reduction in Mps1 T490 autophosphorylation, both the half-life and stable population of Mps1 increase at metaphase kinetochores (Figure 2E,F). However, in the absence of PP1-87B, Mps1 displays a faster dynamic exchange at aligned kinetochores, as expected from enhanced kinase activity (Figure 2E,F). Collectively, these results demonstrate that PP1-87B activity is required to represses the activating autophosphorylation of Mps1 that remains associated with metaphase kinetochores.

PP1-87B/ PP1-γ dephosphorylates Mps1 T-loop

To evaluate whether PP1 directly antagonizes Mps1 T-loop activating autophosphorylation, we started by assessing in vitro the capacity of PP1-γ to dephosphorylate the kinase T-loop. Simultaneous incubation of PP1-γ with recombinant versions of human (Mps1/TTK) and Drosophila Mps1 orthologues led to a concentration-dependent decrease of T676 (T676Ph) and T490 (T490Ph) autophosphorylation, respectively (Figure 3A,C). Moreover, PP1-γ was able to dephosphorylate the T-loop of Mps1/TTK that had been previously activated by autophosphorylation (Figure 3B). In contrast, λ-phosphatase failed to antagonize to the same extent the autophosphorylation of human Mps1/TTK T676 and of Drosophila Mps1 T490 (Figure 3C and Figure 3—figure supplement 1), indicating a higher specificity of PP1-γ towards Mps1 T-loop in vitro.

PP1-87B/PP1-γ dephosphorylates Mps1 T-loop in vitro and in vivo.

(A,B) Western blot analysis of Mps1/TTK T676 autophosphorylation. Recombinant human Mps1 was incubated during 30 min with increasing concentrations of recombinant human PP1-γ (A) or PP1-γ (42.7 nM) was added to previously active Mps1 and phosphorylation of Mps1 T676 (Mps1T676Ph) monitored over time (B). (C) Western blot analysis of Drosophila Mps1 T490 autophosphorylation. Recombinant Drosophila Mps1 was incubated during 30 min with 10 nM of recombinant human PP1-γ or λ-PP and phosphorylation of Mps1 T490 (Mps1T490Ph) monitored. (D) Clustal W alignments of amino acid residues of indicated Mps1 orthologues. Red boxes highlight putative PP1-docking motifs identified in silico by the Eukaryotic Linear Motif (ELM) resource. (E) MBP-PP1-87B pull-downs from total cell lysates of S2 cells expressing EGFP-Mps1WT or EGFP-Mps1K231A/F234A or EGFP-Mps1WT upon depletion of Sds22. Immobilized MBP was used as negative control. Input, beads and flow-through (FT) were probed by western blotting for the indicated proteins. (F) Quantification of Mps1 binding to PP1-87B from the pull-downs in (E). The chemiluminescence signal intensity of EGFP-Mps1 was determined relative to the signal of MBP-PP1-87B beads. The graph represents the quantification of relative levels of EGFP-Mps1 in MBP-PP1-87B pull-downs from at least two independent experiments. The values obtained for EGFP-Mps1WT from control cells were set to 1. (G) MBP-87B pull-downs of purified recombinant Mps1 N-terminus region (104–330 amino acids) harboring the wild-type (6xHis-N-Mps1WT) or mutated PP1-docking motif (6xHis-N-Mps1K231A/F234A). Immobilized MBP was used as negative control. Input, beads and flow-through (FT) were probed by western blotting for the indicated proteins. (H) Western blot analysis of EGFP-Mps1WT and EGFP-Mps1K231A/F234A T-loop dephosphorylation by PP1-γ. EGFP-Mps1WT and EGFP-Mps1K231A/F234A were immunoprecipitated from mitotic S2 lysates and incubated with ATP for 30 min to allow Mps1 T-loop autophosphorylation. Increasing concentrations of recombinant GST-PP1-γ were subsequently added to the reaction mixture and Mps1 T490 phosphorylation (Mps1T490Ph) assessed after 30 min. Immunoprecipitates were probed by immunoblotting (IB) for the indicated proteins. (I) Quantification of Mps1T490Ph levels from the dephosphorylation assay in (H). The chemiluminescence signal intensity of Mps1T490Ph was determined relative to the corresponding signal of EGFP-Mps1 immunoprecipitates. The values obtained for each control reaction were set to 100%. The graph represents the quantification of Mps1T490Ph relative levels from two independent experiments. (J–L) Representative immunofluorescence images (J,K) and corresponding quantifications (L) of Mps1 T490 phosphorylation (Mps1T490Ph) at kinetochores of unaligned (J) and aligned (K) chromosomes from S2 cells expressing EGFP-Mps1WT or EGFP-Mps1K231A/F234A and from PP1-87B depleted S2 cells expressing EGFP-Mps1WT. EGFP-Mps1 transgenes were expressed under the control of Mps1 cis-regulatory region (Althoff et al., 2012). Mps1T490Ph fluorescence intensities were determined relative to EGFP-Mps1 signal (N ≥ 150 unaligned kinetochores from at least 8 cells for each condition and N ≥ 111 aligned kinetochores from at least 7 cells for each condition). Scale bar: 5 μm. Data information: in (F), (I) and (L) data are presented as mean ± SD. Asterisks indicate that differences between mean ranks are statistically significant, ****p<0.0001 (Kruskal-Wallis, Dunn’s multiple comparison test). Numerical source data for this figure are provided in Figure 3—source data 1.

We then sought to examine whether PP1 directly dephosphorylates Mps1 T-loop in a cellular context and determine its implications for SAC silencing. Because PP1 has numerous mitotic substrates acting on a multitude of pathways, its depletion results in pleiotropic phenotypes that preclude a clear and direct analysis of individual events. To circumvent this, we devised a strategy to specifically prevent Mps1 dephosphorylation without impacting the remaining PP1-87B cellular activity. The association of PP1 with the majority of its interactors is mediated by short motifs that bind to the phosphatase hydrophobic groove (Egloff et al., 1997; Wakula et al., 2003). Drosophila Mps1 N-terminus contains a clear match for a PP1-docking motif of the [R/K]VxF – type (Figure 3D). To assess the relevance of Mps1 KVLF231-234 for the interaction with PP1-87B, we converted K231 and F234 to alanine and expressed both the wild type (EGFP-Mps1WT) and mutant versions (EGFP-Mps1K231A/F234A) of Mps1 in S2 cells. The interaction between Mps1 and PP1-87B was confirmed in pull-down assays in which EGFP-Mps1WT from S2 cell lysates bound efficiently to recombinant MBP-PP1-87B but not to the MBP used as negative control (Figure 3E). Conversely, mutating Mps1 KVLF231-234 to AVLA231-234 markedly decreased the ability of EGFP-Mps1 to interact with MBP-PP1-87B, hence validating KVLF as a PP1-binding motif (Figure 3E,F). To demonstrate that PP1-87B directly interacts with Mps1 via the KVLF motif, we performed pull-down assays with MBP-PP1-87B and bacterially purified fragments of Mps1 N-terminus (104–330 amino acids) harboring the wild-type (6xHis-N-Mps1WT) or mutated PP1-docking motif (6xHis-N-Mps1K231A/F234A). While 6xHis-N-Mps1WT bound efficiently to MBP-PP1-87B, the fragment 6xHis-N-Mps1K231A/F234A did not (Figure 3G), thus confirming that the KVLF motif mediates a direct interaction between Mps1 N-terminus and PP1-87B. In accordance, in vitro assays with EGFP-Mps1 immunoprecipitated from mitotic S2 cell lysates showed that recombinant PP1-γ antagonized with discernible less efficiency the T-loop autophosphorylation of EGFP-Mps1K231A/F234A in comparison to EGFP-Mps1WT (Figure 3H,I). Collectively, these results indicate that Mps1 T-loop is a direct substrate of PP1-87B/PP1-γ. However, we found that MBP-PP1-87B failed to pull-down EGFP-Mps1WT from S2 cells depleted of Sds22/PPP1R7 (Figure 3E,F), which suggests that in addition to the KVLF motif on the kinase N-terminus, the Sds22/PPP1R7 regulatory subunit is also required for competent binding of full-length Mps1 to PP1-87B in cell extracts.

To analyse the impact of diminished PP1-87B binding on cellular Mps1 activation and SAC signaling, we monitored Mps1 T-loop phosphorylation and mitotic progression in S2 cells expressing EGFP-Mps1WT and EGFP-Mps1K231A/F234A under control of Mps1 cis-regulatory region (Figure 3—figure supplement 2). Phosphorylation of T490 at unaligned kinetochores was significantly higher for EGFP-Mps1K231A/F234A relative to EGFP-Mps1WT (Figure 3J,L). A similar increment in the T-loop activation status was observed for EGFP-Mps1WT of prometaphase cells depleted of PP1-87B (Figure 3J,L). Notably, the mutation KVLF231-234 prevented the dephosphorylation of Mps1 T-loop also at bioriented kinetochores, mimicking the result obtained in metaphase cells expressing EGFP-Mps1WT in the absence of PP1-87B (Figure 3K,L). As expected from increased T490 autophosphorylation, EGFP-Mps1K231A/F234A exhibits faster dynamics than EGFP-Mps1WT at aligned kinetochores (Figure 3—figure supplement 3A,B). From these results, we conclude that PP1-87B directly controls Mps1 T-loop dephosphorylation at kinetochores during prometaphase and metaphase.

Live-cell imaging revealed that expression of EGFP-Mps1K231A/F234A caused a pronounced metaphase delay when compared to EGFP-Mps1WT expressing cells (Figure 4A,B; Videos 7 and 8). Depletion of BubR1 prevented this delay showing it is SAC-dependent (Figure 4A,B and Video 9). However, the delay is not caused by unstable kinetochore-microtubule interactions, since the inter-kinetochore distances monitored throughout metaphase and the attachments of bioriented kinetochores to cold-resistant microtubule bundles in EGFP-Mps1K231A/F234A cells were similar to those observed in metaphase cells expressing the wild-type kinase (Figure 4C,D and Figure 4—figure supplement 1A). Nevertheless, cells expressing EGFP-Mps1K231A/F234A retained higher levels of Mad1 at aligned kinetochores (Figure 4E,F and Figure 4—figure supplement 1B,C), which is consistent with lasting Mps1 activation and SAC signaling in metaphase. These results strongly suggest that impairing PP1-mediated dephosphorylation of the residual pool of Mps1 that remains associated with kinetochores of bioriented chromosomes is sufficient to delay SAC silencing and prevent anaphase onset despite normal chromosome attachment and high inter-kinetochore tension.

To further confirm the increment in Mps1 activity in EGFP-Mps1K231A/F234A and its impact on SAC signaling, we treated cells with colchicine to generate unattached kinetochores and monitored their capacity to arrest in mitosis. As expected for SAC competent cells, this led to an evident increase in the mitotic index of both EGFP-Mps1WT and EGFP-Mps1K231A/F234A expressing cells (Figure 4G). Consistent with the role of Aurora B in potentiating Mps1 activation and SAC signaling, inhibition of the former with increasing concentrations of Binucleine 2 (Smurnyy et al., 2010) weakened the SAC in EGFP-Mps1WT cells, as revealed by the gradual reduction in the mitotic index of colchicine incubated cultures (Figure 4G). In accordance, Aurora B inhibition negatively affected Mps1 T-loop autophosphorylation and Mad1 accumulation at unattached kinetochores. Strikingly, expression of EGFP-Mps1K231A/F234A partially rescued Mps1 T490 phosphorylation, Mad1 kinetochore recruitment and consistently attenuated the decline of SAC function following Aurora B inhibition (Figure 4G–I). Collectively, these results demonstrate that compromising Mps1 T-loop dephosphorylation is sufficient to delay SAC silencing and mitotic exit. Thus, we conclude that PP1-mediated dephosphorylation of Mps1 T-loop is required for efficient Mps1 inactivation and consequently for prompt SAC silencing.

(A,B) Representative immunofluorescence images (A) and corresponding quantification (B) of Mps1T490Ph levels in the nucleoplasm of control and CENP-C depleted cells in prophase. Mean values for control cells were set to 100% (N ≥ 40 cells for each condition from four independent experiments). Scale bar: 5 μm. (C,D) mitotic progression (C) and mitotic timing (D) of CENP-C depleted S2 cells expressing mCherry-Tubulin and EGFP-Mps1WT, EGFP-Mps1K231A/F234A or EGFP-Mps1K231A/F234A/T490A under control of Mps1 cis-regulatory region. Mitotic progression was monitored through time-lapse microscopy and the mitotic timing was defined by the length of time between nuclear envelope breakdown (NEB) and anaphase onset (AO) (N ≥ 11 cells for each condition from at least two independent experiments). Scale bar: 5 μm. Data information: in (B) and (D) data are presented as mean ± SD. Asterisks indicate that differences between mean ranks are statistically significant, *p<0.05 (Mann-Whitney U test). Numerical source data for this figure are provided in Figure 5—source data 1.

Regulation of PP1 activity during mitosis

The activity of PP1 must be tightly regulated to allow robust Mps1 activation during early mitosis and to ensure its inactivation and SAC silencing when all kinetochores become stably attached to spindle microtubules. Immunofluorescence analysis of S2 cells revealed that PP1-87B preferentially accumulates on kinetochores when chromosomes are bioriented at the metaphase plate (Figure 6A,B). Kinetochore recruitment of PP1 in human cells is mediated in part by SILK and RVSF motifs present on KNL1 N-terminus. Aurora B-dependent phosphorylation of both motifs prevents the binding of PP1 during prometaphase (Liu et al., 2010). As Drosophila Spc105/KNL1 also contains a RVSF motif in its N-terminus, we examined whether a similar mechanism operates in S2 cells to control PP1-87B kinetochore localization. Cultured cells were incubated with Aurora B inhibitor Binucleine two in the presence of colchicine and of the proteasome inhibitor MG132, which respectively generate unattached kinetochores and prevent mitotic exit resulting from SAC abrogation. Inhibition of Aurora B led to an increase of PP1-87B levels at unattached kinetochores (Figure 6C,D), suggesting that similarly to the described for human cells, Aurora B activity limits the association of PP1-87B with unattached/unaligned kinetochores in flies. In addition to the control exerted by Aurora B on PP1 kinetochore localization, CDK1 phosphorylation of PP1 C-terminus was shown to repress the phosphatase activity during early mitosis (Dohadwala et al., 1994; Yamano et al., 1994; Wu et al., 2009; Grallert et al., 2015). In accordance, inhibition of CDK1 with RO-3306 in colchicine-treated S2 cells led to a reduction of Mps1 T490 autophosphorylation at unattached kinetochores, which was partially restored upon depletion of PP1-87B (Figure 6E,F). Thus, although CDK1 might also directly potentiate Mps1 activity (Morin et al., 2012), these results suggest that CDK1-dependent inhibition of PP1-87B contributes to maintain Mps1 active in prometaphase S2 cells. Declining Cyclin B levels were proposed to relieve PP1 inhibition, thereby ensuring high phosphatase activity in late mitosis (Dohadwala et al., 1994; Wu et al., 2009; Grallert et al., 2015). We monitored by live-cell imaging the levels of EGFP-Cyclin B throughout mitosis in S2 cells and as expected found that it starts to be rapidly degraded upon biorientation of all chromosomes (Figure 6G,H and Video 14). Interestingly, we could further observe that proteolysis of Cyclin B also occurs during prometaphase but at a significant lower rate (Figure 6G,H and Video 14). It was recently proposed that a small reduction in CDK1/Cyclin B activity is sufficient to allow PP1 auto-reactivation, which consequently triggers a feedback loop that ensures robust phosphatase activity (Grallert et al., 2015; Rogers et al., 2016). Thus, the modest decline in Cyclin B that takes place before metaphase might be critical to permit PP1-mediated inactivation of Mps1 and SAC silencing.

Discussion

How the SAC is rapidly extinguished allowing anaphase to proceed when the last chromosome becomes bioriented has remained a key unanswered question. The present study shows that efficient SAC silencing requires the inactivation of its most upstream regulator, Mps1 kinase. We demonstrate in vitro and in vivo that PP1 directly antagonizes Mps1 T-loop autophosphorylation rendering the kinase inactive at metaphase kinetochores and in the cytoplasm. In Drosophila this relies on a [R/K]VxF motif that is present on Mps1 N-terminus and requires the PP1 regulatory subunit Sds22. Importantly, Mps1 orthologues encompass several other PP1-binding motifs within different regions of the kinase, such as SILK signatures in the C-terminus of human and mouse Mps1. This suggests that despite the diversification in type and localization of PP1-docking motifs among Mps1 orthologues, the dephosphorylation of the kinase T-loop by PP1 is likely to be conserved.

The calponin homology (CH) domains of Ndc80 and Nuf2 directly mediate Mps1 accumulation at unattached kinetochores, which is proposed to promote the kinase activation required for efficient SAC signaling (Kemmler et al., 2009; Santaguida et al., 2011; Saurin et al., 2011; Nijenhuis et al., 2013). Because microtubules bind to the same surface of the CH domains that interacts with Mps1, microtubules forming end-on attachments outcompete Mps1 for Ndc80 binding (Hiruma et al., 2015; Ji et al., 2015). This competition mechanism is consistent with the reduction of Mps1 from metaphase kinetochores, but is however insufficient to dislodge all Mps1 molecules. As in human and budding yeast (Vázquez-Novelle and Petronczki, 2010; Aravamudhan et al., 2015), Drosophila kinetochores retain a fraction of Mps1 even after the formation of end-on attachments that exert proper kinetochore tension. Intriguingly, although low levels of Mps1 were shown to be sufficient to delay anaphase onset (Herriott et al., 2012; Foijer et al., 2014), the residual pool of Mps1 at metaphase kinetochores does not prevent SAC silencing. Work in budding yeast proposes that this is due to changes on kinetochore architecture imposed by end-on attachments. Stably bound microtubules exert mechanical tension on kinetochores, which increases the distance between the CH domains of the Ndc80 complex and the MELT motifs on Spc105/KNL1, thus preventing phosphorylation of the latter by Mps1 and allowing SAC silencing (Aravamudhan et al., 2015). However, since in higher eukaryotes kinetochore-localized Mps1 dynamically exchanges with the cytoplasmic pool (Jelluma et al., 2010), free diffusion of Mps1 is expected to overcome the physical separation between the kinase receptor and its substrates at metaphase kinetochores. Moreover, the recruitment of Mad1-Mad2 to budding yeast kinetochores occurs exclusively through a linear pathway in which Mps1-phosphorylated MELTs function as docking sites for the binding of Bub1-Bub3 complexes, which in turn act as a loading platform for Mad1-Mad2 localization (London and Biggins, 2014). In Drosophila and in human cells however, an Spc105/KNL1- and Bub1-independent pathway also recruits Mad1-Mad2 to unattached kinetochores (Schittenhelm et al., 2009; Caldas et al., 2015; Silió et al., 2015). This relies on the localization of the RZZ complex at kinetochores, which is itself regulated by Mps1 activity (Santaguida et al., 2010) and is sufficient to activate the SAC in response to unattached kinetochores, even in the absence of KNL1 (Schittenhelm et al., 2009; Silió et al., 2015). Hence, in addition to the physical separation between the Ndc80 complex and Spc105/KNL1 phosphodomain, other mechanisms must operate in metazoans to limit the activity of kinetochore-associated Mps1 and extinguish SAC signaling in metaphase. Here we demonstrate that this is accomplished by PP1-mediated dephosphorylation of Mps1 T-loop. We found that depletion of PP1-87B or preventing its interaction with Mps1 increases the activation status of the residual fraction of Mps1 that persists at kinetochores of bioriented chromosomes. This correlates with higher levels of Mad1 retained on kinetochores and a prolonged metaphase delay that is caused by chronic SAC engagement despite stable end-on attachments and establishment of kinetochore tension. Thus, we reason that intra-kinetochore tension or alterations on kinetochore architecture, which also correlate with SAC satisfaction in Drosophila and human cells (Maresca and Salmon, 2009; Uchida et al., 2009), are important not only to disrupt the interaction of Mps1 with its kinetochore substrates but also to increase the recruitment of PP1 to metaphase chromosomes. Similarly to human cells (Liu et al., 2010), kinetochore levels of PP1-87B in Drosophila S2 cells are significantly higher at bioriented chromosomes. The association of PP1-γ with human kinetochores is directly mediated by SILK and [R/K]VxF motifs on Spc105/KNL1 N-terminus and by the Spindle- and Kinetochore-Associated (Ska) complex (Sivakumar et al., 2016). At tensionless kinetochores, the recruitment of PP1-γ is limited due to phosphorylation of Spc105/KNL1 N-terminus and of the Ska complex by Aurora B, which respectively inhibits the binding of PP1-γ to Spc105/KNL1 and Ska kinetochore localization (Liu et al., 2010; Chan et al., 2012). As microtubules bind to kinetochores and tension is established, the spatial repositioning of the KMN network decreases Aurora-B mediated phosphorylation of the Ska complex and of Spc105/KNL1 N-terminus (Welburn et al., 2010), hence allowing bulk accumulation of PP1-γ, which then catalyses the dephosphorylation of Mps1 T-loop. This provides a mechanism to ensure the complete inactivation of any vestigial Mps1 residing at microtubule-attached kinetochores during late stages of mitosis.

In addition to the MCC generated at unattached kinetochores in prometaphase, APC/C inhibitory complexes are also assembled at the nuclear pore and in the cytoplasm during late interphase and prophase (Sudakin et al., 2001; Lopes et al., 2005; Lince-Faria et al., 2009; Maciejowski et al., 2010; Schweizer et al., 2013; Rodriguez-Bravo et al., 2014). This pre-mitotic MCC defines the minimum length of time a cell will spend in mitosis as it ensures APC/C inhibition until newly assembled kinetochores are able to generate an efficient SAC response. Interestingly, inhibition of Mps1 causes a substantial decrease in pre-mitotic MCC levels and a consequent reduction in Cyclin B levels already in G2 and prophase (Maciejowski et al., 2010; Rodriguez-Bravo et al., 2014). Moreover, a truncated form of Mps1 unable to localize at kinetochores failed to promote the recruitment of Bub1 to unattached kinetochores but was nevertheless sufficient to sustain the assembly of mitotic APC/C inhibitory complexes and delay anaphase onset (Maciejowski et al., 2010). In addition, cytosolic Mps1 activity was shown to be critical to maintain the metaphase arrest caused by Mad1 constitutively tethered to stably attached bioriented kinetochores that were stripped from Mps1 (Maldonado and Kapoor, 2011). These observations indicate that the activity of soluble Mps1 is required to generate the MCC and/or prevent its disassembly during mitosis, further strengthening the idea that removing the kinase from kinetochores is not sufficient for efficient SAC silencing. Therefore, determining how cytosolic Mps1 is inactivated after the SAC signal is extinguished at stably attached kinetochores becomes essential to understand the mechanism that allows timely mitotic exit. Here we show that PP1 directly controls Mps1 activation, not only at kinetochores but also in the cytoplasm. We demonstrate that preventing PP1-mediated dephosphorylation of soluble Mps1 maintains the kinase active in the cytosol and that this is sufficient to significantly delay mitotic exit even in the absence of kinetochore-generated MCC. Although kinetochore recruitment of Mps1 substantially increases the cell capacity to sustain a prolonged mitotic arrest when challenged with spindle poisons (Nijenhuis et al., 2013), during an unperturbed mitosis in which MCC production at kinetochores is minimal (Collin et al., 2013; Dick and Gerlich, 2013), controlled activity of soluble Mps1 is expected to be particularly relevant. We envisage that the inactivation of cytosolic Mps1 is critical to ensure SAC responsiveness at kinetochores during late mitosis so that transition to anaphase occurs without delay after the last unattached kinetochore forms stable attachments.

CDK1 phosphorylation of PP1 C-terminus represses the phosphatase activity during early mitosis (Dohadwala et al., 1994; Yamano et al., 1994; Wu et al., 2009; Grallert et al., 2015). Our data suggest that CDK1-dependent inhibition of PP1-87B contributes to maintain Mps1 active in prometaphase S2 cells. Declining Cyclin B levels were shown to relieve PP1 inhibition (Dohadwala et al., 1994; Wu et al., 2009; Grallert et al., 2015). However, several lines of evidence indicate that the SAC is switched off under condition of high CDK1/Cyclin B activity (Mirchenko and Uhlmann, 2010; Oliveira et al., 2010; Kamenz and Hauf, 2014; Rattani et al., 2014; Vázquez-Novelle et al., 2014), hence challenging the impact that CDK1/Cyclin B-mediated regulation of PP1 might have on SAC silencing. Interestingly, recent mathematical modelling support that small decreases in CDK1/Cyclin B activity are sufficient to initiate PP1 re-activation of and trigger a positive feedback loop that ensures robust phosphatase activity (Rogers et al., 2016). Therefore, slow Cyclin B proteolysis occurring before metaphase in S2 cells might be sufficient to allow PP1-mediated inactivation of Mps1 and SAC silencing when biorientation of all chromosomes is achieved. Another possibility is that global repression of PP1 during early mitosis is ensured by CDK1/Cyclin A. Cyclin A is progressively degraded during prometaphase and its abundance governs the overall stability of kinetochore-microtubule attachments, which increases in metaphase when cyclin A levels fall below a critical threshold (Kabeche and Compton, 2013). It would be interesting to investigate whether decreased CDK1/Cyclin A coordinates this switch in attachment stability at the prometaphase to metaphase transition with the re-activation of PP1 and consequently with SAC silencing. This could explain how PP1 switches off the SAC despite elevated Cyclin B levels in early metaphase.

Taking our results together with previously reported work (Dohadwala et al., 1994; Yamano et al., 1994; Liu et al., 2010; Mochida and Hunt, 2012; Grallert et al., 2015; Qian et al., 2015; Rogers et al., 2016), we propose a model that controls Mps1 activation in a timely manner and perfectly coordinated with mitotic progression. Elevated CDK1 and Aurora B activities during prometaphase, respectively repress PP1 activity and avert the phosphatase localization at unattached kinetochores. This ensures high levels of active kinetochore- and cytosolic Mps1 and therefore, robust SAC signaling in early mitosis (Figure 7). As kinetochores become stably attached to spindle microtubules, Mps1 levels drastically decrease whereas PP1 accumulation increases. This occurs concomitantly with a drop in Cyclin B (and Cyclin A) that allows prompt PP1-mediated inactivation of Mps1 at kinetochores and in the cytoplasm (Figure 7). We propose that this coincides with the dephosphorylation of Mps1 substrates, with the removal of other SAC proteins and with modifications on kinetochore organization. As Mps1 promotes MCC assembly through multi-target phosphorylations along the SAC signaling cascade, the checkpoint is highly responsive to changes in Mps1 activity (Faesen et al., 2017; Ji et al., 2017). Inactivation of Mps1 by PP1 prevents the kinase from counteracting the dephosphorylation of its substrates, thus avoiding futile cycles and ensuring efficient and rapid SAC silencing. This guarantees a swift anaphase onset following chromosome biorientation.

A model for regulation of Mps1 activation/inactivation in mitosis.

Proposed model for regulation of soluble- and kinetochore-Mps1 activation in mitosis. During prometaphase, Aurora B potentiates the recruitment of Mps1 to unattached kinetochores and phosphorylates KNL1/Spc105 to limit PP1 kinetochore association. This allows the accumulation of active Mps1 at unattached kinetochores to instate efficient MCC assembly. In parallel, high levels of active CDK1 repress PP1 activity in the cytoplasm, which prevents the dephosphorylation and consequently inactivation of soluble Mps1. Active Mps1 in the cytoplasm promotes the assembly and/or prevents the disassembly of APC/C inhibitory complexes through mechanisms that are yet to be described. Cytoplasmic-and kinetochore generated MCC orchestrated by Mps1 cooperate to ensure efficient APC/C inhibition during unperturbed early mitosis. End-on attachment of microtubules mediated by the Ndc80 complex and KNL1 prevent the recruitment of Mps1 and exert tension across kinetochores and/or impose alterations on kinetochore architecture. Under these conditions, phosphorylation of KNL1 by Aurora B is minimal, and PP1 becomes enriched at bioriented kinetochores, where it dephosphorylates and inactivates the remaining residual pool of Mps1. Declining levels of active CDK1 allow PP1 to auto-activate and repress Mps1 activity in the cytoplasm, hence ensuring efficient SAC silencing and prompt anaphase onset.

Materials and methods

S2 cell cultures, RNAi and drug treatments

Drosophila S2 cell cultures (S2-DGRC), RNAi synthesis and RNAi treatments were performed as previously described (Conde et al., 2013). At selected time points, cells were collected and processed for immunofluorescence, time-lapse microscopy or immunoblotting. When required, cells were subjected to several drug treatments before being collected and processed for the desired analysis. In order to promote microtubule depolymerisation, cells were incubated with 30 µM colchicine (Sigma–Aldrich, St. Louis, MO) for 2–12 hr. To inhibit Aurora B activity, selected concentrations of Binucleine 2 (Sigma-Aldrich) were added to cultured cells for 2 hr. To inhibit CDK1/Cyclin B activity, 10 µM RO-3306 (Sigma-Aldrich) were added to cultured cells for 1 hr. When required the 20 µM MG132 (Calbiochem, San Diego, CA) was added to cultured cells to inhibit the proteasome. The Drosophila S2-DGRC cell line (stock#6) was acquired from the Drosophila Genomics Resource Center, Indiana University and was not independently authenticated. The cell lines were routinely tested negative for mycoplasma contamination.

Constructs and S2 cell transfection

The pCaSpeR4 harboring EGFP-Mps1WT under control of Mps1 cis-regulatory region was a gift from Christian Lehner (Althoff et al., 2012). Constructs EGFP-Mps1K231A/F234A and EGFP-Mps1K231A/F234A/T490A were generated by site-directed mutagenesis with primers harboring the desired mutation. PCR reactions were performed with Phusion polymerase (New England Biolabs, Ipswich, MA) and pCaSpeR4-EGFP-Mps1WT as template. PCR products were digested with DpnI restriction enzyme (New England Biolabs), used to transform competent bacteria and selected for positives. To overexpress EGFP-Mps1WT and EGFP-Mps1K231A/F234A in S2 cells, Mps1 coding sequence was cloned in frame with N-terminal EGFP under regulation of a metallothionein promoter in the pMT-EGFP-C vector (Invitrogen, Carlsbad, CA) as previously described (Conde et al., 2013). pMT-EGFP-Mps1K231A/F234A was generated by site-directed mutagenesis with primers harboring the desired mutation. PCR reactions were performed with Phusion polymerase (New England Biolabs) and pMT-EGFP-Mps1WT as template. PCR products were digested with DpnI restriction enzyme (New England Biolabs), used to transform competent bacteria and selected for positives. Mad1-EGFP construct under regulation of Mad1 native promoter cloned in the pMT backbone (Invitrogen) was a gift from Thomas Maresca (University of Massachusetts Amherst). The constructs H2B-GFP, mCherry-α-Tubulin and EGFP-Cyclin B have been previously described (Conde et al., 2013). Stable cell lines expressing the indicated constructs were obtained by cotransfection with the pCoBlast and selection in medium with 20 µg/ml blasticidin. Transfections of recombinant plasmids and pCoBlast into S2 cells were performed using Effectene Transfection Reagent (Qiagen), according to the manufacturer’s instructions.

S2 cell lysates, pull-downs, immunoprecipitation and western blotting

For preparation of S2 cell lysates used in pull-down and immunoprecipitation experiments, S2 cultured cells overexpressing EGFP-Mps1WT and EGFP-Mps1K231A/F234A were treated with colchicine for 12 hr. Overexpression of EGFP-Mps1WT and EGFP-Mps1K231A/F234A was induced with 0.1 mM CuSO4 during 12 hr. Cells were harvested through centrifugation at 5000 rpm for 10 min at 4°C and afterwards washed with 2 mL PBS supplemented with protease inhibitors cocktail (Roche, Basel, Switzerland). Cell pellet was resuspended in lysis buffer (150 mM KCl, 75 mM HEPES, pH 7.5, 1.5 mM EGTA, 1.5 mM MgCl2, 15% glycerol, 0.1% NP-40, 1× protease inhibitors cocktail (Roche) and 1× phosphatase inhibitors cocktail 3 (Sigma)) before disruption through freezing in liquid nitrogen. Cell lysates were then clarified through centrifugation at 8000 rpm for 10 min at 4°C and quantified by Bradford protein assay (Bio-Rad). EGFP-Mps1WT and EGFP-Mps1K231A/F234A were immunoprecipitated from 0.5 mg of total cell lysates using the GFP-Trap_MA system (Chromotec GmbH, Planegg-Martinsried, Germany) according to the manufacturer’s instructions. For pull-down assays using S2 cell protein extracts, 0.5 mg of total cell lysates were diluted in a final volume of 500 μl of column buffer (250 mM NaCl, 20 mM Tris-HCl pH 7.4, 1 mM EDTA, 1 mM DTT, 0.05% Tween-20, 1× protease inhibitors cocktail (Roche) and 1× phosphatase inhibitors cocktail 3 (Sigma-Aldrich)) and incubated with 5 μl of MBP or MBP-PP1-87B bound to amylose magnetic beads (New England Biolabs) for 90 min at room temperature with rotation. The magnetic beads and bound protein fraction were collected and washed 3 times with 750 μl of column buffer. For pull-down assays with purified recombinant proteins, 5 μl of 6xHis-N-Mps1WT or 6xHis-N-Mps1K231AF234A were incubated with 3 μl of MBP or MBP-PP1-87B bound to amylose magnetic beads (New England Biolabs) in a final volume of 30 μl of column buffer (250 mM NaCl, 20 mM Tris-HCl pH 7.4, 1 mM EDTA, 1 mM DTT) for 60 min at room temperature with rotation. The magnetic beads and bound protein fraction were collected and washed 3 times with 100 μl of column buffer. Magnetic beads and bound protein were resuspended in Laemmli sample buffer and boiled for 5 min at 95°C. After removal of the magnetic beads, samples were resolved by SDS-PAGE and probed for proteins of interest through western blotting. For western blot analysis, resolved proteins were transferred to a nitrocellulose membrane, using the iBlot Dry Blotting System (Invitrogen) according to the manufacturer’s instructions. Transferred proteins were confirmed by Ponceau staining (0.25% Ponceau S in 40% methanol and 15% acetic acid). The membrane was blocked for 3 hr at room temperature with 5% dry milk in PBS-T. All the primary and secondary antibodies were diluted in PBS-T containing 1% dry milk and the membranes were incubated overnight at 4°C under agitation, then washed three times 10 min with PBS-T and immediately incubated with secondary antibodies for 1 hr at room temperature under agitation. Secondary antibodies conjugated to Horseradish peroxidase (Amersham) were used according to the manufacturer´s instructions. Blots were developed with ECL Chemiluminescent Detection System (Amersham) according to manufacturer’s protocol and detected on X-ray film (Fuji Medical X-Ray Film).

Immunofluorescence analysis

For immunofluorescence analysis of S2 cells, 105 cells were centrifuged onto slides for 5 min, at 1500 rpm (Cytospin 2, Shandon), and simultaneously fixed and extracted in 3.7% formaldehyde (Sigma-Aldrich), 0.5% Triton X-100 in PBS for 10 min followed by thee washing steps in PBS-T (PBS with 0.05% Tween 20) for 5 min each. Immunostaining was performed as described previously (Conde et al., 2013). Images were collected in Leica TCS SP5 II laser scanning confocal microscope (Leica Microsystems, Germany). Data stacks were analyzed and projected using ImageJ software (http://rsb.info.nih.gov/ij/). For immunofluorescence quantification, the mean pixel intensity was obtained from maximum projected raw images acquired with fixed exposure acquisition settings. For kinetochore proteins, the mean fluorescence intensity was quantified for individual kinetochores, selected manually by CID or Spc105 staining. The size of the region of interest (ROI) was predefined so that each single kinetochore could fit into. After subtraction of background intensities, estimated from regions of the cell with no kinetochores, the intensity was determined relative to CID or Spc105 reference and averaged over multiple kinetochores.

Live cell imaging

Live analysis of mitosis was done in S2 cell lines expressing the indicated constructs. 4D datasets were collected at 25°C with a spinning disc confocal system (Revolution; Andor) equipped with an electron multiplying charge-coupled device camera (iXonEM+; Andor) and a CSU-22 unit (Yokogawa) based on an inverted microscope (IX81; Olympus). Two laser lines (488 and 561 nm) were used for near-simultaneous excitation of EGFP and mCherry. The system was driven by iQ software (Andor). Time-lapse imaging of z stacks with 0.8 μm steps covering the entire volume of the cell were collected and image sequence analysis and video assembly done with ImageJ and iQ software. For quantification of EGFP-Mad1 fluorescence at kinetochores, the mean intensity was calculated within the area corresponding to kinetochores and corrected for cytosolic signal. The changes in fluorescence intensity with time were plotted as normalized signal relative to the signal measured at NEB.

Inducible speckle imaging (ISI) was used to measure spindle microtubule turnover in Drosophila S2 cells as previously described (Pereira et al., 2016). Speckle patterns were imprinted using ∼200- to 500-ms-long pulses with a bleaching strength (γ) between 1.5 and 2, a regime which maximizes speckle fluorescence amplitude. A rectangular ROI was defined enclosing a significant portion of the spindle area. Intensity contrast was then determined at each time point of the acquired sequence after the subtraction of a mean dark reference level from each frame. The microtubule turnover half-time was calculated by fitting a double-exponential curve (y0 +A1*exp[−(t − t0)/τ1]+A2 *exp[−(t − t0)/τ2]) to the mean intensity squared-contrast from independent cells for each time point. Before averaging, contrast curves were normalized to a 0-to-1 contrast change defining the pre- and post-ISI transition.

FRAP experiments

FRAP experiments were performed in S2 cells expressing EGFP-MPS1WT, EGFP-MPS1KD or EGFP-MPS1K231A/F234A. When required, colchicine was added to cultures at least 2 hr prior FRAP experiment. Datasets were collected at 25°C with a Leica TCS SP5II scanning confocal microscope (Leica Microsystems, Germany). The EGFP tag was excited using the 488 nm laser line set to 10% and bleached with the 405 nm laser line. Single kinetochores were bleached for three iterations once the EGFP fluorescence signal had become stable. Fluorescence intensity in the bleached area was acquired every 537 ms, 555 ms or 777 ms before and after bleaching. FRAP data analysis was performed with ImageJ. For each measurement the average fluorescence intensity in the bleached area was corrected for background and the ratio to average fluorescence in the cytoplasm determined. Average values before bleaching were set to 100%. The exponential kinetics of FRAP were analyzed by calculating the normalized unrecovered fluorescence at each time point (Finf-F(t))/(Finf-F(0)) where Finf is the value reached at the plateau, F(0) is the value observed in the first frame after bleaching and F(t) is the value at a given time point. FRAP kinetics parameters were determined by one phase exponential association curve fitting to normalized data using GraphPad Prism software.

Fly stocks

All fly stocks were obtained from Bloomington Stock Center (Indiana, USA), unless stated otherwise. The mps1 mutant allele aldG4422 has been described before (Conde et al., 2013). insc-GAL4 was used to drive the expression of UAS-PP1-87BRNAi in neuroblasts from brains of 3rd instar larvae brains. w1118 was used as wild-type control. Fly stocks harboring gEGFP-MPS1WT and gEGFP-MPS1KD (D478A) under control of Mps1 cis-regulatory region were kindly provided by Christian Lehner (Althoff et al., 2012).

Statistical and kinetics analysis

Supplementary information

The phospho-antibody that recognizes human Mps1 T676 autophosphorylation can be used to detect the conserved activating T-loop autophosphorylation on Drosophila Mps1 (T490). The phospho-antibody generated against human Mps1 T676 autophosphorylation readily detected in immunoblots the autophosphorylation of a recombinant wild type version of Drosophila Mps1 (His6-Mps1WT) contrasting with the absence of evident phosphorylation on a catalytically inactive version of the kinase (His6-Mps1KD) (Figure 2—figure supplement 2B). Moreover, immunofluorescence analyses of 3rd instar larval neuroblasts revealed a significant decrease in Mps1 T490 phosphorylation at kinetochores of aldG4422 strong hypomorphic Mps1 mutants (Figure 2—figure supplement 2C,D). Importantly, a similar reduction was observed following the expression of kinase dead EGFP-Mps1 in aldG4422 background when compared with mutants expressing the Mps1 wild-type transgene (Figure 2—figure supplement 2E,F). Taken together, these data validate the phospho-antibody as a read-out for the activation status of Drosophila Mps1.

Decision letter

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Protein Phosphatase 1 inactivates Mps1 to ensure efficient Spindle Assembly Checkpoint silencing" for consideration by eLife. Your article has been favorably evaluated by Anna Akhmanova (Senior Editor) and three reviewers, one of whom, Andrea Musacchio (Reviewer #1), is a member of our Board of Reviewing Editors.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

In this manuscript, Moura, Osswald et al. investigated an important unresolved problem in the spindle assembly checkpoint (SAC) field: how the wait-anaphase signal is inactivated in a timely manner once the SAC has been satisfied. Specifically, the manuscript describes a new facet of the complex regulation of the SAC kinase Mps1. The authors contend that depletion of the Drosophila melanogaster (dm) Protein Phosphatase 1 (PP1) ortholog PP1-87B leads to a mitotic arrest in metaphase, and that this correlates in turn with the presence of the active, phosphorylated form of Mps1 on the kinetochores of aligned metaphase chromosomes, for all apparent levels of tension (using inter-kinetochore distance as its surrogate marker). This is an important observation, because current models based on two 2015 papers in Science by the Kops and Yu laboratories have proposed that microtubule binding prevents kinetochore recruitment of Mps1 through direct competition. At face value, the authors' observations appear inconsistent with this previously proposed model. The authors also make the compelling point that Drosophila cells are the ideal model system to analyze this SAC regulatory mechanism because Dm Spc105/KNL1 MELT-like motifs are not subjected to phospho-regulation, yet PP1 is required for mitotic exit, pointing to Mps1 as a possible crucial target. Indeed, the authors focus on a possible physical interaction of PP1 with Mps1, which ultimately results in the dephosphorylation of the T-loop of Mps1 in metaphase, ensuring timely SAC inactivation. The authors contend that the expression of an Mps1 mutant that escapes this interaction largely phenocopies the depletion of PP1. In addition, the authors show that regulation by Mps1 is not only affecting the kinetochore pool of the kinase, but also extends to the cytosolic form. The authors suggest that this regulation may not be limited to Drosophila Mps1, and that it could be more general.

All three reviewers recognise the significant interest and potential importance of the work. However, they also agree that the data provided do not unequivocally show that Mps1 is a direct substrate of PP1. Main points of criticism are:

1) In Drosophila the PP1-87B isoform accounts for about three quarters of all PP1 activity. Hence, the depletion of PP1-87B will increase the phosphorylation of hundreds of substrates and affect nearly every cellular process. Some of the observed effects may even be mediated by other phosphatases (e.g. PP2A), which are known to be regulated by PP1. Therefore, an increased phosphorylation of Mps1 by the depletion of PP1-87B can at best be used as preliminary evidence for an (in)direct role of PP1 in the dephosphorylation and inactivation of Mps1.

2) One way to delineate the exact contribution of PP1 to Mps1 dephosphorylation is by interference with the involved PP1-targeting subunit. The authors conclude that PP1 directly interacts with Mps1 via a canonical RVxF motif. However, the data are not conclusive. First, the consensus binding motif for PP1 is not conserved in other organisms (Figure 3D) and it is not clear whether this motif resides in an intrinsically disordered region of Mps1, which is a requirement for binding to PP1. Second, RVxF and SILK consensus sequences are present in a third of all proteins and their mere presence does not imply that they also function as PP1 binding motifs. In fact, most consensus PP1 binding sequences do not interact with PP1, often because they reside in structured domains. Most importantly, a PP1 binding mutant of Mps1 still binds nearly normal amounts of PP1 (Figure 3E) and it is therefore not clear how to explain the effects of the expression of this Mps1 mutant. All three reviewers raised this point, which is therefore critical. At present, it cannot be concluded that the increased phosphorylation of Mps1 at its T-loop following mutation of the RVxF sequence is due to a loss of associated PP1. This also implies that the molecular mechanism underlying the metaphase delay effects after expression of the Mps1 K231A/F234A mutant remains unaccounted for.

Essential revisions:

1) All reviewers felt that the pulldowns in Figure 3E are not convincing. It definitely looks like the amount of MBP-PP187B is significantly higher in the WT extracts. The levels of EGFP to MBP were ratioed, but the data are not strongly convincing, as the EGFP looks quite comparable between the WT and mutant. The cell-based data on this mutant are more convincing than the results obtained in vitro. However, with the present data, one of the paper's main hypotheses appears unsubstantiated, and the reviewers recommend a more detailed characterisation of the Mps1-PP1 interaction.

2) The depletion of SDS22 mimicked PP1-87B RNAi (Figure 2—figure supplement 3). This interesting observation was not explored further. In view of the criticisms raised in point 1, it is possible that SDS22 mediates the binding of PP1-87B by Mps1. This possibility could be tested relatively easily.

3) The purified catalytic subunit of PP1 will dephosphorylate nearly every phosphoprotein when added in sufficient amounts, which makes the experiments shown in Figures 3A-C inconclusive. It would have been much more informative to compare the dephosphorylation of Mps1-WT and Mps1-K231A/F234A by limiting concentrations of PP1. If Mps1 directly recruits PP1 via its putative RVxF motif (but see point above) the WT protein can be expected to be dephosphorylated at lower PP1 concentrations than the K231A/F234A mutant.

4) Figure 3—figure supplement 1 is taken as evidence that the λ phosphatase does not dephosphorylate Mps1 at T676. However, panel C shows much faster dephosphorylation with λ phosphatase than with PP1. Likewise, in Figure 3C, the major band disappears with λ phosphatase. The phosphatase units should be clearly defined: a comparison based upon units only makes sense if they are defined in the same manner.

5) A central finding is that PP1 depletion leads to delay in SAC satisfaction due to excess active p-Mps1 at kinetochores (and in the cytosol). Mad1 is also retained at these kinetochores. A possible explanation for this phenotype is that elevated aurora B kinase, as a consequence of PP1 depletion, simply generates unattached kinetochores that retain Mad1 and delay the cell in metaphase. Key in this regard is to convincingly demonstrate that fully stable attachments are formed. To strengthen this conclusion, the authors should better quantify the cold stability data. All that is shown for these experiments are a few images of cold-treated cells.

6) Related to point 5, turnover measurements after photo-activation have been used more commonly, and the authors should elaborate on the validity and robustness of the ISI protocol. Has the ISI approach been validated by comparison to PA measurements? In Figure 1H, the slow turnover component corresponding to KT-MTs has a relatively short half-life in comparison to those reported in previous studies (extending to several minutes). What may be the reason for this? If PA measurements could be considered, they would strengthen this line of enquiry, which is critical for the study. A supplemental video of a representative ISI experiment would be helpful.

Furthermore, the authors may consider the following two comments:

7) The pMps1-T490P seems to localize to the inner centromere rather than the kinetochore in the w1118 larval neuroblasts and the EGFP-WT Mps1 in ald/ald neuroblasts. A CID internal (inner centromere?) Mps1-T490P staining pattern is also observed in Figure 6C. Can the authors share their thoughts on the relevance of Mps1 localization patterns? For example, are there multiple Mps1 sub-populations: cytosolic, kinetochore, inner centromere?

8) Figure 6A-F should be discussed in the Results section, not in the Discussion. The authors speculate that the phosphorylation of PP1 by Cdk1 (at the C-terminus, not the N-terminus as indicated in the fourth paragraph of the Discussion) contributes to keeping Mps1 active at the beginning of mitosis. However, there are no data supporting the notion that Mps1-associated PP1-87B is actually phosphorylated by Cdk1 at the beginning of mitosis. More generally, regulation of PP1 by Cdk1 is unlikely to contribute to the switch from the SAC on to the SAC off condition, as the SAC is switched off before Cyclin B destruction can start, i.e. it is switched off under conditions of high Cdk1 activity. Furthermore, while the focus is on CDK1/CyclinB-mediated regulation of PP1, CDK1/CyclinA is also likely contributing to this pathway during prometaphase. Please add some discussion of the contributions of Cyclin A during prometaphase. Does MG132-treatment block the increase in kinetochore-associated PP1 at aligned kinetochores that is nicely shown in Figures 6A, B?

Author response

Essential revisions:

1) All reviewers felt that the pulldowns in Figure 3E are not convincing. It definitely looks like the amount of MBP-PP187B is significantly higher in the WT extracts. The levels of EGFP to MBP were ratioed, but the data are not strongly convincing, as the EGFP looks quite comparable between the WT and mutant. The cell-based data on this mutant are more convincing than the results obtained in vitro. However, with the present data, one of the paper's main hypotheses appears unsubstantiated, and the reviewers recommend a more detailed characterisation of the Mps1-PP1 interaction.

We agree with the reviewers and following their recommendation we performed a more detailed characterization of the interaction between Mps1 and PP1. We repeated the pull-downs, but this time under more stringent conditions (pull-down buffer supplemented with 250 mM NaCl + Tween 0.05%). This allowed us to clearly show that the interaction between Mps1 and PP1-87B is severely compromised when the KVLF motif is mutated to AVLA (Mps1K231A/F234A). We have presented these data and its corresponding quantification in Figure 3E, F of the revised manuscript.

To demonstrate direct binding of Mps1 to PP1-87B and the requirement of the KVLF motif for such an association we produced recombinant fragments of Mps1 N-terminus (104-330 aa) harboring the KVLF motif and its mutated version AVLA. MBP-PP1-87B was able to pull-down the recombinant Mps1WT N-terminus as opposed to the Mps1K231A/F234A fragment. These results demonstrate a direct interaction between Mps1 N-terminus and PP1-87B that is mediated by a canonical RVxF motif. We have included these new data in Figure 3G of the revised manuscript.

2) The depletion of SDS22 mimicked PP1-87B RNAi (Figure 2—figure supplement 3). This interesting observation was not explored further. In view of the criticisms raised in point 1, it is possible that SDS22 mediates the binding of PP1-87B by Mps1. This possibility could be tested relatively easily.

We agree that the increase of Mps1 T-loop phosphorylation upon Sds22 depletion is an interesting observation. Following the reviewers’ suggestion, we assessed whether Sds22 might be necessary for the interaction between Mps1 and PP1-87B. For that, we performed pull-downs using MBP-PP1-87B and lysates from S2 cells expressing EGFP-Mps1WT and depleted of Sds22. Binding of EGFP-Mps1WT to MBP-PP1-87B is readily detected when lysates of control cells are used. Notably, the absence of Sds22 decreased this interaction. These results are consistent with a role for Sds22 in mediating the interaction between PP1-87B and full-length Mps1 and have been included in Figure 3E, F of the revised manuscript. Thus, although Mps1 N-terminus is able to bind PP1-87B in vitro through a canonical RVxF motif (please see response to point 1), the interaction of PP1-87B with full-length Mps1 from cell lysates seems to require the involvement of SdS22 regulatory subunit.

3) The purified catalytic subunit of PP1 will dephosphorylate nearly every phosphoprotein when added in sufficient amounts, which makes the experiments shown in Figures 3A-C inconclusive. It would have been much more informative to compare the dephosphorylation of Mps1-WT and Mps1-K231A/F234A by limiting concentrations of PP1. If Mps1 directly recruits PP1 via its putative RVxF motif (but see point above) the WT protein can be expected to be dephosphorylated at lower PP1 concentrations than the K231A/F234A mutant.

We thank the reviewers for this important suggestion. To address this, we immunoprecipitated EGFP-Mps1WT and EGFP-Mps1K231A/F234A from lysates of S2 cells depleted of endogenous PP1-87B. Immunoprecipitated Mps1 was incubated with ATP and with increasing concentrations of PP1. We found that the T-loop of Mps1WT was dephosphorylated at lower PP1 concentrations than the Mps1K231A/F234A mutant, thus validating KVLF as a PP1-docking motif and Mps1 as a direct substrate of PP1. We have presented these data on Figure 3H, I of the revised manuscript.

4) Figure 3—figure supplement 1 is taken as evidence that the λ phosphatase does not dephosphorylate Mps1 at T676. However, panel C shows much faster dephosphorylation with λ phosphatase than with PP1. Likewise, in Figure 3C, the major band disappears with λ phosphatase. The phosphatase units should be clearly defined: a comparison based upon units only makes sense if they are defined in the same manner.

The panel C on Figure 3—figure supplement 1 of the original manuscript does not depict the dephosphorylation of Mps1 T-loop but instead the dephosphorylation of an artificial substrate by λ phosphatase and by PP1. As described in the corresponding figure legend, the graph represented the dephosphorylation of DiFMUP over time in an EnzChek phosphatase assay. This control experiment was performed to demonstrate that although at the indicated units of λ phosphatase failed to efficiently dephosphorylate Mps1-T loop the phosphatase was nevertheless active towards other phospho-substrate. We apologize for not making it clear and to avoid misinterpretation we decided to remove these data from the revised version of the manuscript.

We agree with the reviewers that the comparison between PP1 and λ phosphatase based upon units only makes sense when these are defined in the same manner. Therefore, we repeated most of the experiments using PP1 and λ phosphatase at the same molar concentration, which is unambiguous and allows valid comparisons to be made. The results obtained confirmed a higher specificity of PP1 towards Mps1 T-loop and are presented in Figure 3A-C and Figure 3—figure supplement 1 of the revised manuscript.

5) A central finding is that PP1 depletion leads to delay in SAC satisfaction due to excess active p-Mps1 at kinetochores (and in the cytosol). Mad1 is also retained at these kinetochores. A possible explanation for this phenotype is that elevated aurora B kinase, as a consequence of PP1 depletion, simply generates unattached kinetochores that retain Mad1 and delay the cell in metaphase. Key in this regard is to convincingly demonstrate that fully stable attachments are formed. To strengthen this conclusion, the authors should better quantify the cold stability data. All that is shown for these experiments are a few images of cold-treated cells.

We agree with the reviewers. Following their suggestion, we quantified the% of kinetochores bound to cold-resistant microtubule bundles in control, PP1-87B- and Ndc80-depleted cells, as well as in cells expressing EGFP-Mps1WT, EGFP-Mps1K231A/F234A. The results from the quantifications are depicted in Figures 1G and Figure 4—figure supplement 1A of the revised manuscript and clearly demonstrate that fully stable attachments are formed upon depletion of PP1-87B or expression of Mps1K231A/F234A.

6) Related to point 5, turnover measurements after photo-activation have been used more commonly, and the authors should elaborate on the validity and robustness of the ISI protocol. Has the ISI approach been validated by comparison to PA measurements? In Figure 1H, the slow turnover component corresponding to KT-MTs has a relatively short half-life in comparison to those reported in previous studies (extending to several minutes). What may be the reason for this? If PA measurements could be considered, they would strengthen this line of enquiry, which is critical for the study. A supplemental video of a representative ISI experiment would be helpful.

We used ISI to measure microtubule turnover rates because this method is a robust alternative to ROI based techniques, where marking of tubulin is typically done by imaging a mask onto the sample to trigger the photoswitch, generating a pattern confined to the objective’s depth of focus. Although the pattern is constrained to the focal plane, out-of-focus planes still experience a (homogeneous) photoswitch, precluding sequential z-stack imprinting to generate a 3D pattern. In ISI, the fluorescence pattern after the pulse is 3D, hence unrelated to the particular focal plane at the time of the imprinting and thus covers the whole spindle with measurable speckle spots. Thus, different z-layers can be chosen after acquisition. This contributes to operational robustness and operational simplicity when compared to conventional ROI-switch techniques. Not the least, bright speckles decay and dark speckles recovery are globally and mutually measured through a statistical measure (intensity contrast), insensitive to global, acquisition-related, bleaching.

For a detailed description of ISI please consider the article by Pereira and colleagues (2016) [PMID: 26783303], where ISI has also been used to measure spindle microtubule turnover dynamics in Drosophila S2 cells. The half-life value determined by Pereira et al. (2016) for the slow turnover component in metaphase was 21 seconds, which is very similar to the reference values obtained in previous studies using photobleaching: Buster et al. (2007) [PMID: 17553931] and Goshima et al. (2008) [PMID: 18443220] respectively reported half-life values of ~ 20 seconds and 30 seconds for the stable population of metaphase microtubules in S2 cells. Moreover, a fly stock available in the lab (UASp-alphaTub84B.tdEOS) allowed us to determine during the revision period of the manuscript the turnover rates of spindle microtubule in metaphase Drosophila neuroblasts by photoconversion (~ 40 seconds). Thus, the ISI measurements described in our manuscript (~ 40-45 seconds) and in Pereira et al. (2016) for kinetochore-microtubules of Drosophila metaphase cells are comparable to the values obtained when photobleaching or photoconversion approaches are used. Collectively, these data support the validity and robustness of the ISI protocol as a method to measure microtubule turnover rates. Following the reviewers’ recommendation, we have provided supplemental videos of representative ISI experiments in the revised version of the manuscript (Videos 4 and 5).

Since we resorted to ISI to benefit from its abovementioned advantages over the conventional ROI-based methods and the values that we obtained are very similar to the reference values reported for S2 cells, performing PA measurements to confirm the stability of kinetochore-microtubule interactions as suggested by the reviewers would only cover the purpose of using the ISI technique. Note that the ISI data is also supported by interkinetochore measurements and cold-stability quantifications that show fully stable attachments in PP1-87B depleted cells.

Furthermore, the authors may consider the following two comments:

7) The pMps1-T490P seems to localize to the inner centromere rather than the kinetochore in the w1118 larval neuroblasts and the EGFP-WT Mps1 in ald/ald neuroblasts. A CID internal (inner centromere?) Mps1-T490P staining pattern is also observed in Figure 6C. Can the authors share their thoughts on the relevance of Mps1 localization patterns? For example, are there multiple Mps1 sub-populations: cytosolic, kinetochore, inner centromere?

We do not have an explanation for the inner-centromeric localization of Mps1-T490Ph observed in larval neuroblasts. One can speculate on the existence of an inner centromeric subpopulation of active Mps1 that is more evident in fly neuroblasts. However, at this point, we do not have solid data to confirm this notion. In S2 cells however, the phospho-antibody consistently stains the kinetochore. The inset on the bottom panel of Figure 6C of the original manuscript is misrepresenting the actual localization pattern. The inset depicts the magnification of two kinetochores from two different adjacent kinetochore pairs, thus wrongly suggesting a CID internal Mps1-T490Ph signal. We apologize for this and selected an appropriate pair of kinetochores from the same image, which is now presented on Figure 6E of the revised manuscript.

8) Figure 6A-F should be discussed in the Results section, not in the Discussion. The authors speculate that the phosphorylation of PP1 by Cdk1 (at the C-terminus, not the N-terminus as indicated in the fourth paragraph of the Discussion) contributes to keeping Mps1 active at the beginning of mitosis. However, there are no data supporting the notion that Mps1-associated PP1-87B is actually phosphorylated by Cdk1 at the beginning of mitosis. More generally, regulation of PP1 by Cdk1 is unlikely to contribute to the switch from the SAC on to the SAC off condition, as the SAC is switched off before Cyclin B destruction can start, i.e. it is switched off under conditions of high Cdk1 activity. Furthermore, while the focus is on CDK1/CyclinB-mediated regulation of PP1, CDK1/CyclinA is also likely contributing to this pathway during prometaphase. Please add some discussion of the contributions of Cyclin A during prometaphase. Does MG132-treatment block the increase in kinetochore-associated PP1 at aligned kinetochores that is nicely shown in Figures 6A, B?

Following the reviewers’ suggestion we moved the discussion of the data presented in Figure 6A-F of the original manuscript to the Results section under the title “Regulation of PP1 activity during mitosis”. We have also added new data supporting that Aurora B activity limits the accumulation of PP1-87B at unattached/unaligned kinetochores. This is presented in Figure 6C, D of the revised manuscript. Furthermore, to address the reviewers’ concerns, we have provided a more critical discussion about the regulation of PP1 by CDK1/Cyclin B and of its relevance for SAC silencing. A possible contribution of CDK1/Cyclin A in controlling the PP1-mediated transition from the SAC on to the SAC off condition has been added to the Discussion.

European Research Council (PRECISE)

European Research Council (CODECHECK)

Helder Maiato

FLAD Life Science

Helder Maiato

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Geert Kops (UMC, Utrecht, The Netherlands), Thomas Maresca (University of Massachusetts Amherst, USA) and Christian Lehner (University of Zurich, Switzerland) for antibodies, constructs and fly stocks. We thank Carla Sofia Lopes (i3S, IBMC, University of Porto) and all the members of the Sunkel laboratory for critical and helpful discussions. This article is a result of the project Norte-01-0145-FEDER-000029 - Advancing Cancer Research: From basic knowledge to application, supported by Norte Portugal Regional Operational Programme (NORTE 2020), under the PORTUGAL 2020 Partnership Agreement, through the European Regional Development Fund (FEDER). MO is supported by a fellowship from the GABBA PhD program from the University of Porto, PD/BD/105746/2014. JB is supported by an FCT PhD grant SFRH/BD/87871/2012. CC is supported by an FCT investigator position and funding (IF/01755/2014). HM is funded by PRECISE and CODECHECK grants from the European Research Council, FLAD Life Science 2020, and the Louis-Jeantet Young Investigator Career Award.

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